Auxotrophic strains were generated using the SAT1-flipping strategy (Sasse and Morschhauser, 2012). For the deletion of amino acid biosynthetic genes in C. dubliniensis, the SAT1 reusable cassette was PCR amplified from the plasmid pSFS2A with primers containing ∼65 bp of sequence identical to the 5′ and 3′ regions immediately up and downstream of the HIS1, LEU2, and ARG4 open reading frames (ORFs; primer pairs EMO001/EMO002, EMO003/EMO004 and EMO005/EMO006, respectively; Supplementary Table S1). pSFS2A is a plasmid derived from pSFS2 (Reuß et al., 2004) that contains the SAT1 reusable cassette in the backbone of vector pBC SK+ instead of pBluescript II KS and that was kindly provided by Joachim Morschhauser (U. Würzburg). The cassettes were transformed into the CD36 and Wü284 wild type C. dubliniensis strains by electroporation as previously described (Kohler et al., 1997; Staib et al., 2001). For selection of transformants, cells were grown on YEPD medium plates containing 400 μg/ml of nourseothricin (NAT) for 48 h at 30°C. Correct integration of the SAT1 cassette was confirmed by colony PCR of the 5′ and 3′ junctions. For the excision of the SAT1 cassette cells were grown overnight at 30°C in YEP medium containing 2% maltose and then plated on YEPD plates containing 50 μg/ml NAT. After 24 h of growth at 30°C, small colonies were selected and further verified for the loss of NAT resistance by growth on YEPD + 400 μg/ml NAT plates for 48 h at 30°C, as has been described previously (Sasse and Morschhauser, 2012; Porman et al., 2013). Six successive rounds of gene deletion/flipping were needed to obtain the triple auxotrophic mutant. In the final strain, the loss of the three genes was confirmed by PCR with primers that anneal inside the ORF of each gene and by the inability of the strains to grow in synthetic defined (SD) media lacking histidine, leucine or arginine.

For the generation of the equivalent auxotrophic strains in C. tropicalis, we first attempted to use the SAT1-flipping strategy amplifying the deletion cassette with long oligonucleotides that have the 5′ and 3′ homology regions in the same way as for C. dubliniensis. Although we did obtain transformants using this strategy, the integration of the deletion cassette was not in the correct locus. To increase the specificity of the recombination process in this species we used longer homology arms flanking the SAT1 cassette. For this purpose, ∼900 bp regions flanking the HIS1, LEU2, and ARG4 ORFs were PCR amplified from genomic DNA and cloned adjacent to the SAT1 cassette in the pSFS2A or pEM008 plasmids (see below for the description of pEM008 and Supplementary Table S1 for primers used). Preparation of the deletion cassettes for transformation was done either by digesting with the outermost restriction enzymes (see Supplementary Table S1), or by PCR using oligos that anneal to the cloned homology arms as a template. Transformation was performed in the MYA-3404 and AM2005/0093 C. tropicalis strain backgrounds, and selection of transformants and verification of correct integration was done as for C. dubliniensis. Flipping out of the SAT1 marker when using the pSFS2A cassette was achieved by growing C. tropicalis cells for 2 days at 30°C in either liquid or solid YEP medium containing 2% maltose (Porman et al., 2013). Alternatively, flipping out of the markers coming from the pEM008 plasmid involved growing cells overnight at 30°C in either YNB medium with 2% casamino acids (Chauvel et al., 2012) or in synthetic medium containing 2% succinate (Gerami-Nejad et al., 2004). Cells were subsequently screened in YEPD supplemented with 400 μg/ml NAT plates for the loss of NAT resistance before proceeding to delete the second allele. As for C. dubliniensis, the absence of the gene was also verified by colony PCR with primers that amplify part of the ORF and by the inability of the strains to grow in SD media lacking the corresponding amino acid. The details of all generated strains are provided in Table 1.

To clone the C. albicans HIS1, LEU2, and ARG4 genes for use as nutritional markers these genes were PCR amplified with their corresponding promoter and terminator using genomic DNA from the wild type SC5314 strain (see Supplementary Table S1 for primers). Each gene was cloned in the pCR-BluntII-TOPO vector using the Zero Blunt TOPO PCR cloning kit (Invitrogen) following the protocol described by the manufacturer (see Table 2 for plasmid names). The SAT1 and CaHygB genes were PCR amplified from plasmids pSFS2A (Reuß et al., 2004) and pYM70 (Basso et al., 2010), respectively (see Supplementary Table S1 for primers). These genes were cloned in the pCR2.1-TOPO vector using the TOPO TA Cloning Kit (Invitrogen) using the manufacturer’s protocol. All cloned genes were verified by sequencing and the name and details of each resulting plasmid (including the GeneBank accession numbers) are shown in Table 2.

The overall methodology used to delete genes using the auxotrophic strains and markers described here is the same as that previously described for C. albicans (Figure 2) (Noble and Johnson, 2005; Hernday et al., 2010). Gene deletion cassettes were generated by fusion PCR of three fragments: the sequence coding for the nutritional marker, and ∼350 bp sequences flanking the 5′ and 3′ ends of the ORF to be deleted. The PCR amplification of the HIS1, LEU2, and ARG4 nutritional markers was performed using plasmids pEM001, pEM002, and pEM003 as a template, respectively, with primers EMO126 and EMO622. These primers have a region that anneals to the TOPO vectors where the markers are cloned, and a second region that anneals to the primers used for the amplification of the flanks. A specific 20 bp sequence that functions as a “barcode” can be introduced to facilitate the identification of each gene deletion mutant. These barcode sequences can be included as part of the synthesis of primer EMO622 between the sequences GCAGGGATGCGGCCGCTGAC and AGCTCGGATCCACTAGTAACG as has been described (Noble and Johnson, 2005). PCR amplification of the flanks of the gene to be deleted was performed using C. dubliniensis or C. tropicalis genomic DNA as a template, together with primers designed to amplify the ∼350 bp region upstream and downstream of the ORF. The following two sequences were included at the 5′ end of the reverse primer for the 5′ flank amplification and the forward primer of the 3′ flank amplification, respectively: CACGGCGCGCCTAGCAGCGG and GTCAGCGGCCGCATCCCTGC. These sequences allow annealing of the flanks and the auxotrophic markers amplified with primers EMO126 and EMO622 for the fusion PCR reaction. Fusion PCR was performed as described (Hernday et al., 2010) by mixing the three PCR fragments in one final PCR reaction together with the forward primer used to amplify from the 5′ flank and the reverse primer used to amplify from the 3′ flank.

Generation of the deletion cassettes with the SAT1 and CaHygB drug-resistance markers was performed as for the auxotrophic markers, but using the plasmids pEM025 and pEM021, respectively, as PCR templates. Similarly, the markers for C. tropicalis transformations with the markers from C. maltosa and C. dubliniensis that were originally used for C. albicans were PCR amplified from previously described plasmids (Noble and Johnson, 2005). These plasmids are pSN40 (C. maltosa LEU2), pSN52 (C. dubliniensis HIS1), and pSN69 (C. dubliniensis ARG4).

The products of fusion PCR reactions were purified and concentrated using a QIAGEN MinElute PCR Purification Kit, and between 0.5 and 2.5 μg of DNA were transformed by electroporation as described (Kohler et al., 1997; Staib et al., 2001). Selection of transformants was performed by growing cells on SD medium lacking the corresponding amino acid, or on YEPD plates supplemented with 400 μg/ml NAT or 500 μg/ml hygromycin B. Verification of correct integration into the genome was performed by colony PCR of the 5′ and 3′ junctions. In addition, after deleting the second allele, the absence of the target gene was verified by colony PCR with primers that amplify a region within the target ORF. Re-integration of the C. albicans HIS1, LEU2, and ARG4 nutritional markers in C. dubliniensis and C. tropicalis was performed at the corresponding loci. The fusion PCR protocol described above was used to conjugate the C. albicans nutritional markers with ∼350 bp flanks of the C. dubliniensis and C. tropicalis nutritional genes. The corresponding primers are provided in Supplementary Table S1. Selection of transformants in SD media lacking the corresponding amino acid also provided proof of complementation of the auxotrophies by these markers in C. dubliniensis and C. tropicalis.

In our experiments, the efficiency of recycling of the SAT1 cassette derived from pSFS2A in C. tropicalis cells was much lower than that in C. albicans. To improve the efficiency of SAT1 excision in C. tropicalis, we tested replacing the C. albicans MAL2 promoter with its C. tropicalis homolog, or by using the PCK1 promoter of C. albicans or C. tropicalis (Figure 3). The C. albicans MAL2 promoter was previously cloned using BamHI and SalI sites in the plasmid pSFS2A (Reuß et al., 2004). However, there is a second SalI site in the SAT1 ORF of plasmid pSFS2A that precludes re-using this site. To circumvent this issue, we used PCR fusion to combine a fragment containing the desired promoter and a 359 bp segment of the 5′ end of the SAT1 gene from pSFS2A. The resulting PCR product was then cloned into pSFS2A using the BamHI site and an EcoRV site that is present within SAT1. The primer pairs used to PCR amplify the promoter and the SAT1 fragment are the following: for the C. tropicalis MAL2 promoter, EMO178 – EMO179 and EMO176 – EMO177; for the C. tropicalis PCK1 promoter, EMO197 – EMO210 and EMO177 – EMO211; and for the C. albicans PCK1 promoter, EMO242 – EMO243 and EMO177 – EMO244. For the fusion PCR, the following primer pairs were used: C. tropicalis MAL2 promoter, EMO177 – EMO178; C. tropicalis PCK1 promoter, EMO177 – EMO197; C. albicans PCK1 promoter, EMO177 – EMO242. All primer sequences are shown in Supplementary Table S1. As shown in Table 2, the resulting vectors were named pEM004 (contains C. tropicalis MAL2 promoter), pEM008 (contains C. tropicalis PCK1 promoter), and pEM010 (contains C. albicans PCK1 promoter). Flipping out of the SAT1 cassette when using the C. tropicalis MAL2 promoter was achieved by growing cells overnight at 30°C in YEP medium containing 2% maltose. When using the C. albicans or C. tropicalis PCK1 promoters, cassette flipping was induced using either YNB medium with 2% casamino acids (Chauvel et al., 2012) or synthetic medium containing 2% succinate (Gerami-Nejad et al., 2004). Selection of cells that had lost the SAT1 marker was done by growing isolated colonies in YEPD plates for 24 h and replica plating on YEPD plates supplemented with 400 μg/ml NAT.

To generate a vector that contains the cassette to epitope tag genes with an immunoprecipitable peptide and whose SAT1 marker can be efficiently excised in C. tropicalis, we cloned the sequence coding for a 13x MYC tag into plasmids pEM008 and pEM010. As described above, the SAT1 flippable cassette in these two plasmids uses the PCK1 promoter to express the FLP gene (Figure 3). The 13x MYC tag sequence was amplified from the plasmid pADH34 (Hernday et al., 2010) using primers EMO249 and EMO250. This PCR product, that also contains the FRT site at the 3′ end of the MYC tag, was cloned into the BamH1 site of pEM008 and pEM010 that is immediately upstream of the PCK1 promoter. Thus, induction of the FLP recombinase will excise the SAT1 cassette leaving only the epitope tag at the C-terminus of the gene and the FRT site downstream of it. As shown in Table 2, the resulting plasmids were named pEM018 (C. tropicalis PCK1 promoter) and pEM019 (C. albicans PCK1 promoter).

Tagging of genes using these vectors was performed using a similar procedure to the one described for tagging C. albicans genes with the pADH34 vector (Hernday et al., 2010). For efficient transformations, homology regions for directing homologous recombination reactions were extended as was the case for most genetic modifications in C. tropicalis. We therefore generated ∼350 bp homology regions by fusion PCR of the DNA regions flanking the target gene together with the genetic marker cassette. The cassette was PCR amplified from vectors pEM018 and pEM019 using primers EMO623 and EMO624 (Supplementary Table S1). The primers used to extend the homology flanks were designed to amplify the ∼350 bp immediately up and downstream of the stop codon of the target gene, not including the stop codon. In addition, the reverse complement sequence of primer EMO623 (CCGTTAATTAACCCGGGGATCCG) was added to the 5′ end of the reverse primer to amplify the 5′ flank, and the reverse complement sequence of primer EMO624 (GATCCACTAGTTCTAGAGCGGCCGCC) was added to the 5′ end of the forward primer to amplify the 3′ flank. These two sequences allow the flanks to anneal to the tagging cassette for the fusion PCR. Fusion PCR was performed as before to combine the three PCR fragments, using the forward primer employed to amplify the 5′ flank and the reverse primer used to amplify the 3′ flank. The product of the fusion PCR was purified, concentrated and transformed by electroporation, as described above. Selection of transformants was performed on YEPD plates with 400 μg/ml NAT for 48 h at 30°C, and correct integration verified by colony PCR of junction regions. Flipping out of the SAT1 cassette was performed as described above by growing cells in YNB medium with 2% casamino acids. Verification of gene tagging was performed by PCR amplification and Sanger sequencing of the ORF-tag junction.

The ORFs of mNeonGreen or mScarlet-I fluorophores (Shaner et al., 2013; Bindels et al., 2017) were codon optimized for expression in C. albicans including a GGSG linker on the 5′ end, a stop codon, and flanking XhoI sites¹. These sequences were cloned into pSFS2A which had been digested with XhoI and treated with calf intestinal phosphatase. Correct orientation of the insert was confirmed by restriction digest analysis and Sanger sequencing (Figure 3). Details of the resulting plasmids can be seen in Table 2 including GeneBank accession numbers. To tag EFG1 and WOR1 with the mNeonGreen or mScarlet-I fluorophores, the tagging cassette was amplified from plasmid pSFS2-mNeonGreen or pSFS2-mScarlet using primers with ∼80 bp homology sequences to the region immediately up and down of the corresponding ORF stop codon, excluding the stop codon itself (for primers see Supplementary Table S1). The cassette was then transformed by electroporation as described above and transformants were selected on YEPD + 400 μg/ml NAT plates for 48 h at 30°C. Correct integration of the fluorophore tag was verified by colony PCR of integration junctions. WOR1 was tagged in cells in the white state, which were afterwards switched to opaque as previously described (Mancera et al., 2015). For visualization under the microscope, cells of all tagged strains were grown overnight in liquid Spider medium at room temperature. Cells were imaged using a Zeiss Observer.Z1 equipped with a CoolSnap HQ₂ camera, Colibi 470 nm LED light source and Zeiss filter set 38 (GFP) and 45 (Texas Red).

Candida dubliniensis is a member of the CTG clade that has only been isolated in the diploid form and for which no conventional sexual cycle has been identified. To be able to delete genes in this species, we generated parental strains that are auxotrophic for histidine, leucine and arginine (Table 1). These strains were generated by deleting the open reading frames (ORFs) of both alleles of the HIS1, LEU2, and ARG4 genes employing the SAT1-flipping strategy previously used for gene deletions in C. albicans (Reuß et al., 2004). In these strains, two of the auxotrophies can be used to select for replacement of each allele of a given target gene with the corresponding amino acid biosynthetic gene. The third auxotrophy is convenient as it can be used to select for reintegration of the gene of interest to verify the phenotype of the mutant, as well as other genetic modifications. For the generation of deletion cassettes, the HIS1, LEU2, and ARG4 genes of C. albicans were independently cloned with their endogenous promoters and terminators in the same plasmid backbone so that they can be amplified with a unique set of primers (Table 2). The amino acid sequence identity of the HIS1, LEU2, and ARG4 genes between C. albicans and C. dubliniensis is 98, 97, and 99%, respectively. Despite the similarity of these genes at the amino acid level, the differences at the nucleotide level (90–94% identity) are expected to lower the probability that the amino acid biosynthetic markers will integrate back into the corresponding endogenous loci in C. dubliniensis.

As shown in Figure 2, the strategy presented here employs fusion PCR to join DNA fragments together to generate the deletion cassettes for transformation. PCR amplicons of ∼350 bp flanking the 5′ and 3′ ends of the target gene are fused to the sequence of the desired amino acid biosynthetic gene. The PCR-amplified deletion cassettes are then transformed into auxotrophic parental strains and selection performed in medium lacking the corresponding amino acid. The correct integration of the amino acid marker into the locus of interest is then verified by colony PCR. As a proof of concept, we integrated the C. albicans HIS1, LEU2, and ARG4 genes into C. dubliniensis strains deleted for the corresponding endogenous genes. Importantly, all three C. albicans genes complemented for the auxotrophies in C. dubliniensis. C. dubliniensis strains carrying the three integrated C. albicans genes are also useful as a control to compare other gene deletion mutants using this method. Overall, the gene deletion strategy presented here is very similar to that successfully used to generate large collections of mutants in C. albicans and C. parapsilosis. We have successfully generated several gene deletion mutants in C. dubliniensis that will be described elsewhere (E. M. et al., manuscript in preparation). The two alleles of target genes were successfully deleted by two consecutive transformations, suggesting that the process to knockout these genes or the nutritional markers did not cause extensive aneuploidies, which is a possible side effect of genetically modifying these species.

The C. dubliniensis auxotrophic strains were generated in two different genetic backgrounds, CD36 and Wü284. CD36 is the type strain and is the only C. dubliniensis strain whose genome has been sequenced to date (Jackson et al., 2009). Wü284 on the other hand is the strain that has been most used to study C. dubliniensis at a molecular level. The first genetic modification systems in C. dubliniensis using the URA3 marker were developed in this genetic background, and even the group that reported the sequence of CD36 used Wü284 for some phenotypic assays (Staib et al., 2001; Jackson et al., 2009). We generated the triple auxotrophic strains in both backgrounds as a starting point for gene deletion construction. As shown in Table 1, we also generated single auxotrophic strains for each amino acid and the double histidine/leucine auxotroph. All of these strains are available and will greatly facilitate genetic studies of C. dubliniensis.

As is the case for C. dubliniensis, no conventional sexual cycle or haploid forms are known for C. tropicalis. Thus, the same difficulties exist for genetic manipulation of the species as for other closely related members of the CTG clade. To facilitate genetic modification of C. tropicalis, we generated a set of strains that are auxotrophic for two of the three following amino acids: histidine, leucine and arginine (Table 1). As for C. dubliniensis, these parental strains were generated by deleting the ORFs of both alleles of HIS1, LEU2, and ARG4 using a SAT1-flipping strategy. However, in contrast to strain construction in C. dubliniensis, deletion of the amino acid biosynthetic genes in C. tropicalis utilized longer homology arms for efficient targeting of the cassette into the genome. As described in detail in the Methods, ∼900 bp flanking sequences were cloned into the plasmid containing the SAT1 marker to generate the deletion cassettes for the three genes. The observed differences in the integration efficiency between the two species may reflect dissimilarities in their DNA double-strand break repair mechanisms. For example, non-homologous end-joining (NHEJ) rather than homologous recombination could be the preferred repair pathway in C. tropicalis when the flanking homology regions are short.

The same strategy outlined for C. dubliniensis can be used to delete genes in auxotrophic C. tropicalis strains. The cloned HIS1, LEU2, and ARG4 genes are used as templates for the generation of deletion cassettes which are transformed into the corresponding parental strains. We used either the amino acid biosynthetic genes from C. albicans as described above for C. dubliniensis, or C. maltosa LEU2 and C. dubliniensis HIS1 and ARG4 that have been used for strain construction in C. albicans. The protein sequence identity between C. tropicalis, C. albicans, C. maltosa, and C. dubliniensis amino acid biosynthetic genes ranges between 86 and 99%, and all of the genes tested complemented the corresponding auxotrophy in C. tropicalis. The correct integration of the auxotrophic markers was verified by PCR of the DNA junctions between the deleted gene and the targeting cassette. In addition to the three nutritional markers described above, we also cloned the SAT1 and CaHygB genes into a similar plasmid backbone (Table 2) and used these for providing resistance to nourseothricin and hygromycin B, respectively. These markers can be PCR amplified with the same set of primers used for the nutritional markers to generate gene deletion cassettes. We have successfully used both nutritional markers and the drug resistance markers in different combinations to effectively delete multiple genes in C. tropicalis (Mancera et al., 2015; Anderson et al., 2016; E. M. et al., manuscript in preparation). As with C. dubliniensis, the two alleles of different target genes were successfully deleted in tandem, which suggests that these strains do not contain aneuploid chromosomes.

The C. tropicalis auxotrophic strains were generated in MYA-3404 and AM2005/0093 genetic backgrounds. MYA-3404 is the strain whose genome has been sequenced (Butler et al., 2009), while AM2005/0093 is a strain that was isolated from urine in Colombia and has been phylogenetically placed close to MYA-3404 by multi-locus sequence typing (Jacobsen et al., 2008). However, AM2005/0093 was naturally homozygous for the mating type-like (MTL) locus and thus was one of several strains picked to study the molecular mechanisms that control phenotypic switching, as switching is MTL-regulated in C. albicans (Mancera et al., 2015; Anderson et al., 2016). We had previously reported some of the auxotrophic strains generated in the AM2005/0093 background. Overall, the strains generated in the two genetic backgrounds (Table 1) offer an ideal starting point for the generation of gene deletion mutants in C. tropicalis.

The SAT1 flippable cassette was originally developed for facilitating genetic manipulation of the C. albicans genome (Reuß et al., 2004). It is a powerful genetic engineering tool for diploid species since it allows the deletion of both alleles of a target gene even in the absence of available nutritional markers. Several variations of the system have been constructed, but in general the system employs the FLP site-specific recombinase to act on two FLP recognition target sites (FRT) that are present in direct repeats flanking the cassette containing the selectable marker. The FLP recombinase catalyzes excision of the region between the FRT sites and removes the selection marker, thus allowing for repeated use of the same marker for other genetic modifications. In a recent version of the system, the FLP gene is conditionally expressed using the promoter of the C. albicans MAL2 gene which is induced when cells are grown in media containing maltose as the carbon source (Sasse and Morschhauser, 2012). Overall, the system combines a drug-resistance marker that allows efficient selection of transformants with an effective system for recycling the marker.

The same SAT1-flipping strategy has also been used successfully in C. dubliniensis and C. tropicalis to delete genes and was used here for the generation of auxotrophic strains. Nevertheless, the FLP-mediated excision of the deletion cassette was much less efficient in C. tropicalis than in C. albicans. Even after several days of growth in maltose-containing media very few cells lost the SAT1 marker, hindering the overall utility of this reusable system. The difficulties in SAT1 recycling likely reflect poor induction of FLP using the CaMAL2 promoter in C. tropicalis cells. We therefore first replaced the CaMAL2 promoter with the endogenous version from C. tropicalis, but FLP-induced recombination was still rare. As a second strategy, we tested driving FLP expression from the promoter of the C. tropicalis PCK1 gene or the C. albicans PCK1 gene (Figure 3). The PCK1 promoter has previously been used to regulate gene expression in C. albicans as it is induced by gluconeogenic carbon sources (Gerami-Nejad et al., 2004; Chauvel et al., 2012). We therefore tested SAT1 recycling by growing cells in succinate or casamino acids as the carbon source. Using either the C. albicans or the C. tropicalis PCK1 promoter we found that excision of SAT1 in C. tropicalis was as efficient as that using the MAL2 promoter-driven FLP flipper system in C. albicans. We have therefore engineered an efficient system for marker recycling in C. tropicalis.

The recyclable SAT1 cassette has also been used as part of a construct to epitope tag genes in C. albicans (Hernday et al., 2010). For this purpose, SAT1 is placed adjacent to the epitope tag so that nourseothricin can be used to select transformants, and the FLP gene is then induced to flip-out the SAT1 cassette leaving only the epitope tag fused to the gene and an FRT site downstream of the tag. Removal of the SAT1 cassette diminishes the potential for disruption of the untranslated region (UTR) to impact gene function. To adapt this system for C. tropicalis, we cloned a 13x MYC tag into the plasmids described above that use the C. tropicalis or C. albicans PCK1 promoter to regulate FLP expression (Figure 3). When tested, this strategy allowed efficient excision of the SAT1 marker after epitope tagging several genes. The results of the immunoprecipitation of these genes will be described elsewhere (E. M. et al., manuscript in preparation). It is important to note that genome-wide sequencing data from the immunoprecipitation experiments does not show any evidence of large-scale copy number variations in these strains, again consistent with them having euploid genomes.

It has not been trivial to use conventional fluorophores such as GFP to visualize the cellular localization of gene products in C. tropicalis. For example, although we are able to generate the appropriate genetically modified strains, a number of protein-GFP fusions or GFP reporters were not visible under the microscope, with the exception of highly expressed histone-GFP and histone-mCherry fusions (Porman et al., 2011). The yeast-enhanced GFP and YFP have also been expressed successfully from a strong constitutive promoter when not fused to a target gene (Defosse et al., 2018). To enable cell biological approaches in C. tropicalis, we engineered plasmids that contain the next-generation fluorophores mNeonGreen and mScarlet that had been codon-optimized for C. albicans² (Figure 3). These plasmids can be used to generate gene-fluorophore fusions by amplifying the fluorophore cassette using oligonucleotides with sequence identity to the C-terminus of the gene of interest. The fluorophores were cloned in the SAT1 flippable system so that transformants can be selected using nourseothricin, and the drug-resistance marker flipped out by induction of the FLP gene. As a proof of concept, we tagged the transcription factors Efg1 and Wor1 with mNeonGreen, and also tagged Efg1 with mScarlet. As observed in Figure 3, it was possible to visualize the cellular localization of Efg1 when fused to both fluorophores. Efg1 is a transcription factor that regulates morphological transitions in C. tropicalis including filamentation and white-opaque switching (Mancera et al., 2015). The Wor1-mNeonGreen fusion was also detected, and this protein was only visible in cells that were in the hybrid/opaque state and not visible in cells in the white state. This is in agreement with the role of this transcription factor in determining the cell’s phenotypic state (Porman et al., 2013; Anderson et al., 2016). The discrete localization of both transcription factors in the nucleus is consistent with their function as transcriptional regulators. We have therefore successfully engineered tools that can be used to tag proteins with next-generation fluorophores in order to improve the visualization of gene products in C. tropicalis cells.